Not applicable.
Not applicable.
The present invention is generally directed toward luminescent methods and compositions for measuring various biological events, such as cell death, membrane damage, cell proliferation, or enzyme activities. In these methods, something occurring as a result of enzyme activity is able to produce light, which is detected in a luminometer or other instrument capable of detecting light. The invention is more particularly directed to methods of measuring various biological events, such as cytotoxicity, membrane damage, cell proliferation, enzyme activities, or some combination of these events, by coupling the activities of enzymes, which may be supplied by the investigator, or which may have been released from dead or damaged cells, with production or consumption of high-energy molecules such as adenosine triphosphate (ATP) or nicotinamide adenine dinucleotide (reduced form) (NADH), and subsequently measuring the concentrations of these high-energy molecules by evaluation of the light produced by a light-producing molecule, such as a luciferase.
Assays for cell death and cell proliferation are very widely performed in many areas of biological and clinical research. They may be used to assess the cytotoxic effects of a drug candidate (such toxicity may be either desirable or undesirable), measure the activity of complement, measure programmed cell death (apoptosis), quantify growth-inhibitory or growth-enhancing effects, detect and characterize environmental toxins, determine the sterility or bioburden of a sample, assess drug sensitivity or resistance of a patient""s tumor cells or a culture of an infectious organism, or simply determine cell number. One of the most usefull and efficient applications of cell death and proliferation assays is in high-throughput screening (HTS), a collection of methods currently used by many pharmaceutical and biotechnology companies to determine the properties of large libraries of drug candidates very rapidly. However, the methods of determining cell death and proliferation currently in use all suffer from important limitations. Some of these limitations make the assays impractical for use in HTS, and also limit their utility in traditional research environments.
Assays in current use for cell death, or cytotoxicity assays, fall into several categories. One category is xe2x80x9creleasexe2x80x9d assays, in which a substance released by dying cells is measured. Often the substance is an enzyme, such as lactate dehydrogenase (LDH) or glyceraldehyde-3-phosphate dehydrogenase (G3PDH). Traditional enzyme-release assays have exploited the fact that these enzymes create NADH, which can be observed by UV spectroscopy at 340 nm. An alternative is to couple production of NADH to generation of a colored dye, as in the LDH-based CellTiter(copyright) assays currently available from Promega. However, these processes are slow and lack sensitivity. For example, the current product from Promega recommends seeding of 5,000-100,000 cell per well, depending on the cell type, and an incubation time with the chromogenic reagents of one hour or more. Other enzymes used in this way include phosphatases, transaminases, and argininosuccinate lyase. These enzymes are typically present in low quantities in most cells, and they do not lend themselves to simple activity assays, making the process of determining cell death cumbersome and insensitive.
Another variety of release assay involves pretreatment of the target cells with a radioactive isotope, generally 51Cr or 3H. Upon lysis, the radioactive contents are released and counted in a scintillation counter. Aside from the problems of handling and waste disposal of radioactive materials, these assays also suffer from various artifacts, and are tedious because of the pretreatment and recovery steps required. The same process can also be carried out with fluorescent dyes, such as bis-carboxyethyl-carboxyfluorescein or calcein-AM, but, again, pretreatment is required, and the dyes are spontaneously released at a significant rate by healthy cells.
Another type of release assay is the luminescent assay of ATP released from dead or damaged cells. However, as it is actually used, this is a proliferation assay, and it is discussed further below along with other proliferation assays.
Another category of cytotoxicity assay makes use of dyes which are able to invade dead cells, but not living cells. An example of such a dye is trypan blue. These assays are useful for examining individual cells, but for quantification of overall cytotoxicity they are inefficient because each cell must be counted individually, either by laborious microscopic analysis or by very expensive and time-consuming flow cytometry. Moreover, some modes of death.(such as complement-mediated lysis) are not easily assessed by this method, because the dead cell remains intact for a limited period of time, after which it can no longer be counted because it has disintegrated.
Yet another category of cytotoxicity assays includes those methods directly related to apoptosis. These assays typically look for either protein markers of apoptotic processes or particular effects on DNA that are uniquely associated with apoptosis. The methods are generally slow and tedious, and thus are not suitable for high-throughput screening applications. Another method of studying apoptosis is to look at the ATP:ADP ratios in a cell, which change in a distinct way as the cell enters apoptosis. These assays may be performed by coupled luminescent methods (Bradbury et al. (2000) J. Immunol. Methods 240:79). However, while these methods are useful for qualitative definition of the mode of death, they have no advantages over the ATP-release assay in quantitative determinations of cytotoxicity or proliferation.
Proliferation assays are methods of measuring numbers of live cells. This may be better for some applications than measuring cell death or damage. For example, proliferation assays are able to reveal cytostatic, growth-inhibitory, and growth-enhancing effects which yield no readout in a cytotoxicity assay. Proliferation assays are also in common use as indirect cytotoxicity assays, but there are serious drawbacks with this approach; these are discussed below in connection with the ATP-release assay. Proliferation assays also fall into several categories. Assays of metabolic activity are in widespread use in research laboratories. The commonly used methods make use of tetrazolium salts, which are reduced in living cells to colored formazan dyes. One advantage of these methods is convenience, especially with the newer dyes (MTT and WST-1). The dye is added to the cell culture, and the absorbance of the formazan is read, typically after 0.5-12 hours. However, there are several important disadvantages. Metabolically active cells reduce the dyes at rates much greater than quiescent cells; the readout may therefore be a poor reflection of the cell number. Moreover, the readout is not an instantaneous xe2x80x9csnapshotxe2x80x9d of the quantity of live cells when the measurement is taken, but rather a complex integral of metabolic activity over the preceding time interval, whose mathematical relationship to the actual live cell number involves the half-life of the dye as well as variations in metabolic activity. Metabolism-based assays are not suitable for measurement of cellular cytotoxicity (for example, the activities of cytotoxic T lymphocytes), or any other assay system in which live cells other than the target cells are present, because these other cells will yield a substantial and often ill-defined background signal. Finally, various artifacts have been associated with the use of these dyes (see for example O""Brien et al. (2000) Eur. J. Biochem. 267:5421-5426; Natarajan et al. (2000) Cancer Detection and Prevention 24:405-414). Although they have not been thoroughly characterized with respect to their effects on cell metabolism, it is known that various agents, such as antioxidants, can interfere with performance of the dyes.
Another kind of proliferation assay actually measures the ability of the cells to grow. This is the colony-forming unit (CFU) assay. It is typically used with cells that grow rapidly and are capable of growth from single cells. The cells are diluted and plated on appropriate growth media, and the colonies are counted when they appear. This method is quite accurate, but is extremely tedious and quite expensive. The labor-intensive aspect of this method is exacerbated by the fact that multiple dilutions of each sample must usually be plated in order to ensure that at least one plate will yield a countable number of colonies.
Finally, cytotoxicity assays can be used as proliferation assays (and vice versa). To use a cytotoxicity assay to count live cells, one simply kills all the cells and performs the assay. (In some cases it may be necessary to wash the cells first, because the readout may depend on a molecule that may have been released into the supernatant by cells that have already died.) The most important example of this approach is the ATP-release assay, mentioned above (Crouch et al. (1993) J Immunol. Methods 160:81). Although strictly speaking this is a cytotoxicity assay, in that ATP released by dead cells is measured, it is rarely used as a direct cytotoxicity assay, because of the very short lifetime of extracellular ATP. Instead, the cells are killed with a lytic agent before the ATP is measured by the luciferase reaction. Thus even though the assay is basically a cytotoxicity assay, if it is to be used to measure cytotoxicity, it is an indirect method, like the other proliferation assays. The ATP-release assay has a number of advantages not enjoyed by many other proliferation assays. It is more sensitive, with a limit of detection of 10-100 cells. It is much faster, with completion of the lysis and assay steps in as little as 3 minutes. Because of the sensitivity, relatively low volumes and small numbers of cells are required. It is really the only assay currently on the market that is sufficiently rapid and sensitive for use in HTS. However, important disadvantages should be noted. The ATP content of cells is subject to strong metabolic fluctuations, which will cause artifacts. Moreover, the assay can be performed only a single time, immediately after cell lysis; if that opportunity is somehow missed, the experiment must be repeated. Finally, in cytotoxicity mode, the assay suffers from very important drawbacks that are common to all proliferation assays used in this mode. The initial seeding of the wells or reaction vessels with cells must be very accurate, because the cytotoxicity readout depends on differences (which may be small) between numbers of surviving cells, and any scatter in the initial seeding contributes substantially to the noise in the results. This leads to the second problem, which is that a direct readout is almost always preferable to a signal that depends on subtracting two large numbers, as the user must do to use a proliferation assay to measure cytotoxicity. Another very important difficulty is a time-consuming problem with this approach which does not involve the actual assay step. Typically the user adds a potentially toxic compound or agent, waits for death or damage to occur, and then measures the result. The length of time the user must wait depends on the method. If the user is measuring cell death directly, then it can be measured as soon as it occurs, perhaps within minutes. However, if the user is measuring live cells in order to derive the cytotoxicity signal, then the user must wait much longer, until the cytotoxic effect has had sufficient time to cause a detectable difference between the test sample and the control. Furthermore, the required time interval is not known in advance, and if the experiment is stopped too soon, it must be repeated (or abandoned, since the user will not know whether a result showing no difference between test and control is due to the lack of an effect or insufficient time to show an effect). Thus in an HTS mode, where minutes are critical, there is an intervening step in this process requiring an interval of time which may be anywhere from 10 minutes to several days, and which cannot be predicted in advance. This is a serious drawback to the use of any proliferation assay for cytotoxicity work, including the ATP-release assay.
Another type of viability assay, also luminescent, is represented by xe2x80x9cCytoLite,xe2x80x9d a trade name for a mitochondrion-based viability assay (Woods and Clements (2001) Nature Labscene UK March, 2001, 38-39). This method is homogeneous, but requires a 15-minute incubation, and a further 10-minute xe2x80x9cdark-adjustmentxe2x80x9d period before the luminance read; it is therefore too slow for high-efficiency HTS. It is also a viability assay and is subject to all of the drawbacks mentioned above as inherent to viability and proliferation assays.
A cytotoxicity assay based on release of alkaline phosphatase from target cells of killer lymphocytes was described in 1994 (Kasatori et al. (1994) Rinsho Byori 42:1050-1054). This assay method is not suitable for use with other types of cells in general, since most cells do not express alkaline phosphatase in sufficient quantity. Moreover, it involves the use of a substrate whose general effects on cells have not been characterized. It is not a homogeneous or high-throughput assay.
A luminescent cytotoxicity assay described in a 1997 report is based on stable transfection of target cell lines of interest with luciferase or B-galactosidase (Schafer et al. (1997) J. Immunol. Meth. 204:89-98. In terms of sensitivity, this assay represents an advance over conventional release assays; however, the disadvantages of this approach are serious. First, stable transfection itself is a labor-intensive and expensive procedure; yet this must be done for every target cell line of interest if the method of Schafer et al. is to be used. Stable transfection does not always work, and, if it does, may alter the metabolic characteristics of the target cell and thereby severely complicate interpretation of the results of the experiment. The method may not be applicable to cell types outside of these that may be transfected in this manner: expression systems would be different, and the enzymes might be produced in insufficient quantities, in inactive form, or not at all. Moreover, the assay is not homogeneous. Instead the cell culture supernatant must be separated from remaining live cells prior to running the assay. This in itself is a very serious drawback in the high-throughput screening environment, since it adds a complex step to the procedure. Finally, according to the authors, luciferase had a half-life of approximately 30 minutes under the conditions used, and this was found to be inadequate for quantification of cell death in prolonged assays.
Again in 1997, a coupled luminescent method was published (Corey et al. (1997) J. Immunol. Meth. 207:43). This method addressed several of the problems of all of the above methods. This was a release assay, but with important differences from other release assays. G3PDH activity was measured by coupling its cognate glycolytic reaction to the following reaction in glycolysis, which is carried out by phosphoglycerokinase (PGK). The PGK reaction produced ATP, which was then measured by luciferase, which was provided in a separate cocktail, yielding a luminance signal. The limit of detection was  less than 0.1 cell, which was superior to the sensitivity of any other available assay and adequate for almost any application. The assay was relatively fast (xcx9c12-15 minutes). Since it provided a direct readout of cytotoxicity, it suffered from none of the disadvantages of proliferation assays used in cytotoxicity mode. The luminance signal continued to increase with time, a feature which allowed the user to decide when an acceptable signal had been achieved xe2x80x9con the fly.xe2x80x9d Nevertheless, the GPL assay had its own disadvantages which prevented it from being commercially viable. It was cumbersome to execute, in that it involved four transfer steps (cocktail to reaction vessel, sample to reaction vessel, luciferase to luminance vessel, aliquot of reaction to luciferase) and two incubations prior to the actual read. Moreover, because the assay cocktail was not compatible with live cells, tests involving bacteria, erythrocytes, or other non-adherent cells or microbes were still more tedious, because the live cells had to be separated from the supernatant by centrifugation prior to the assay. Finally, like all the methods described above, the assay could be used in cytotoxicity mode or in proliferation mode (the latter by killing all the cells prior to the readout), but not both, with a single sample. These features contributed to the unsuitability of the GPL assay for use in high-throughput screening, especially the necessity of several transfers and the separation of the cells from the supernatant. It was also of limited utility for research use because of its complexity of operation.
As mentioned above, an important disadvantage shared by most cytotoxicity and proliferation assays currently available is that they do not permit measurement of both cytotoxicity and proliferation in a single sample. Release assays, such as the GPL assay, permit quantification of cell rupture or damage, but do not reveal the presence or amount of live cells present. On the other hand, proliferation assays, such as the MTT and ATP-release assays, allow quantification of live cells, in either a non-destructive (MTT) or destructive (ATP-release) mode, but yield no direct information about the degree of cell death that may have occurred. Ideally, the worker would prefer to obtain these two independent pieces of information from the same sample.
In summary, the cytotoxicity and proliferation assays currently available are far from ideal. The traditional release assays suffer from poor sensitivity and speed. Metabolism-based assays are slow, inaccurate with respect to actual cell number, and subject to serious artifacts. CFU assays are too slow and tedious for routine use. ATP-release assays are destructive, one-time assays of moderate sensitivity, and they have numerous important drawbacks as cytotoxicity assays. Although the published coupled luminescent assay (GPL) is superior to the other cytotoxicity and proliferation assays in many ways, it nevertheless is cumbersome and impractical for use in high-throughput screening or research environments because of the processing, numerous transfer steps, and lack of a dual cytotoxicity/proliferation mode.
Today""s drug-discovery environment involves high-throughput screening of inhibitors or other modulators of enzyme activity. Among the enzymes of great interest are phosphatases, which participate in many vital signaling and metabolic pathways. However, assay methods in current use for phosphatases are burdened with a number of drawbacks, including poor throughput or sensitivity, the use of radioactivity, and difficulty of interpretation due to the use of unnatural substrates and/or reaction conditions. Poor throughput and/or sensitivity are often due to the nature of the assay; for example, assays utilizing antibodies against phosphorylated target molecules generally require extended incubations, assays making use of electrophoretic separations are too slow to allow the throughput desired, and assays using radioactivity are inherently incovenient and also suffer from poor throughput. In particular, fluorescence polarization (FP) assays are currently under consideration for high-throughput procedures in some cases. However, these assays, which generally make use of antibodies or other ligands directed against phosphorylated target molecules for detection of phosphatase activity, generally require long incubation times for ligand-target association that significantly reduce the value of these assays in high-throughput screening. These assays also typically involve multiple additions of antibodies or other ligands, and/or wash steps, as well as the design, synthesis, and subsequent ongoing cost of fluorophore-containing biomolecules or synthetic compounds. There is also the possibility that a molecule under study as a modulator of phosphatase activity will give a false signal by binding the fluorophore itself, by otherwise quenching or enhancing its fluorescence, or by blocking the target site on the phosphorylated protein. Finally, many FP assays, and other assays which rely on detection of a phosphorylated target molecule, suffer from an additional disadvantage in that the phosphatase activity yields a negative signal, i.e., a decrease in the phosphorylated molecule which is the target of detection. Such a negative signal is generally considered inferior to a positive signal in enzymology. For one thing, several kinds of artifacts can give rise to a negative signal, including protease contamination or unexpected denaturation of a critical protein. Moreover, a negative signal is usually limited in its dynamic range by its very nature.
Another class of phosphatase assay strategies is based on detection of phosphate liberated by the enzymatic activity. One possibility is radiolabeling of the phosphate group, which can then be separated and counted in some manner. Although this method is still in use in research, it is extremely inconvenient, involving the expense of the label itself, the difficulty and expense of creating or purchasing the labeled compound, a separation step, and the danger and tedium of dealing with the radioactive products. The primary non-radioactive method of detecting phosphate is the use of the malachite green reaction (Mahuren et al. (2001) Anal. Biochem. 298:241), which is quite slow and involves multiple reaction steps, making it unsuitable for high-throughput applications. Another methods of detecting phosphate, which is a coupled luminescent scheme, is useful in devices for environmental or food sampling (Karube, M. (1998) Japanese Patent Application Number 10121688), but involves multiple mixing steps and the use of immobilized enzymes with flow cells in a portable sampling device, making it unsuitable for a high-throughput screening environment. In any case this method has never been shown to be compatible with phosphatase activities. Moreover the oxidizing agents produced in the detection reaction (including hydrogen peroxide) might inactivate a large class of important phosphatases containing active-site thiol groups.
In contrast to the phosphatase assay strategies mentioned above, which can make use of either natural or general peptide/protein substrates, other strategies make use of molecules that are designed more to ease the problem of detection than as ideal substrates for the phosphatase under study. The use of these highly unnatural substrates in high-throughput screening procedures poses a different set of problems, especially problems of interpretation. In most cases the unnatural substrate has quite different kinetic parameters from the actual in vivo substrate. The corollary of this is that when inhibitors or modulators of phosphatase activity are found by such procedures, their characteristics in reactions with the actual in vivo substrate may prove to be very different, especially if competitive inhibition is involved. This is even more likely to be the case if the unnatural substrate has a substantially higher Km (Michaelis constant) for the enzyme than the natural substrate, since competitive inhibitors identified in such a system may successfully compete for the weakly binding unnatural substrate, but may be ineffective against the strongly binding, natural substrate. Similarly, important inhibitors may not be identified by such a system, especially if the substrate is smaller, more labile than, or kinetically distinct from the natural substrate. For example, p-nitrophenylphosphate is a commercially important substrate for alkaline phosphatase, because it is very labile and yields a colorimetric result, but its use in inhibitor screening applications could lead to false, rejection of good inhibitors. An inhibitor might be strong enough to exhibit useful inhibition of the natural reaction, but not strong enough to prevent most of this very labile ester from being hydrolyzed. Similarly, the inhibitor might block the active site in such a way that the natural reaction is prevented, but small molecules such as p-nitrophenylphosphate, phenacyl phosphate, luciferin phosphate, or 1,2 dioxetanes (see below) can still enter the active site and be hydrolyzed. This could lead to rejection of valuable xe2x80x9chitsxe2x80x9d in a screening situation. In short, when the reaction being studied is not the same as the natural reaction that is the desired target, there is a substantial risk that the information gathered will not be biologically useful or relevant.
A luminescent phosphatase assay has been reported that employs a 1,2 dioxetane as a substrate (Adam et al. (1996) Analyst 121:1527; Olesen et al. (2000) Methods Enzymol. 326:175). A related method employs a substrate that leads to generation of a dioxetane in situ (Catalani et al. (1999) Analytica Chimica Acta 402:99). A third method employes phenacyl phosphate as the substrate, followed by reaction with lucigenin (Sasamoto et al. (1995) Anal. Chim. Acta 306:161). These methods work only with alkaline phosphatases, and are not readily extensible to other phosphatases, since a new substrate and/or reaction series might have to be designed and synthesized for each phosphatase. In many or most cases this may be impossible or prohibitively expensive. Alkaline phosphatases typically have very different substrate specificities from the protein phosphatases that are of greatest interest in today""s biology, such as protein tyrosine phosphatases and serine/threonine phosphatases. Moreover, the methods are not rapid, homogeneous assays; for example, the assay recently reported by Olesen et al. involves 3-4 transfers and at least 2 separate incubations, over a period of at least 30 minutes. This would make it most inconvenient for a high-throughput setting. Another serious drawback of these approaches, discussed above, is the use of unnatural substrates.
Another molecule that has been used as a substrate in phosphatase assays is luciferin phosphate (Mountfort et al. (1999) Toxicon 37:909; Miska and Geiger (1988) Biol. Chem. Hoppe-Seyler 369:407). The principle of the assay is that generation of free luciferin by hydrolysis of luciferin phosphate (catalyzed by the phosphatase) may lead to light production in a reaction that contains luciferase and ATP, but a limiting amount of luciferin. In the 1988 work alkaline phosphate was used, but in the 1999 work, luciferin phosphate was used in an assay of protein phosphatase 2A. In both cited references the assay was slow (30-60 minutes for the enzymatic-reaction step alone), and non-homogeneous (involving at least one transfer after initiation). While it is interesting that protein phosphatase 2A hydrolyzes this highly unnatural substrate, the rate of hydrolysis was so poor that the detection limit was more than 1000-fold worse than by fluorimetric methods (however, these fluorimetric methods also required one hour, involved multiple steps, and required highly unnatural substrates). While it is unknown whether this work can be transferred to other protein phosphatases, it is clear that such hypothetical methods, if possible, would likely be insensitive, very slow, and non-homogeneous, and would also make use of unnatural substrates, with all the disadvantages discussed above.
Accordingly, there is a need in the art for assays that are practical for use in high-throughput screening. The present invention fulfills this need and further provides other related advantages.
Briefly stated, the present invention provides a variety of coupled luminescent methods and compositions for use in various assays, including for assaying cytotoxicity, membrane damage, cell proliferation, and enzymatic activity. Luminescent methods have an important advantage over other liquid-phase methods in that the sensitivity of luminescent detection of most phenomena is greater than the sensitivity of any other method. For example, electrochemiluminescent (ECL) analysis of Western blots is now the gold standard in sensitivity, and ECL methods are the most sensitive in enzyme immunoassays. xe2x80x9cCoupled luminescentxe2x80x9d methods are methods in which the activity of the enzyme or enzymes of interest is xe2x80x9ccoupledxe2x80x9d in some manner to production or consumption of a high-energy molecule, such as ATP or NADH, which is a luminescent substrate for one or more of the biological luciferases. Luciferases are enzymes which produce light as they consume such high-energy molecules. Properly designed coupled luminescent assays are able to combine the advantages of specific assays for enzyme function with the very great sensitivity of luminescent detection methods. In these systems the inherent sensitivity of luciferase detection is enhanced by the xe2x80x9camplificationxe2x80x9d effect of enzyme turnover, which produces thousands, millions, or billions of high-energy molecules for each molecule of enzyme.
In one embodiment of the present invention, the measurement takes place in a one-step xe2x80x9chomogeneousxe2x80x9d system; a homogeneous system is one in which the sample is mixed with the reagent cocktail, and no separations or further transfers are required prior to readout. The enzyme or enzymes whose activity is being measured (in enzymatic activity mode) or the enzyme or enzymes released from cells (in cytotoxicity, membrane-damage, proliferation, or combined cytotoxicity/proliferation mode) are coupled in a single reaction vessel to production of ATP, NADH, or another high-energy molecule which is a substrate for a luciferase; the luciferase then produces light from the chemical energy of the high-energy molecule. The increase or decrease in the luminance signal is related to the concentration(s) of the enzyme or enzymes whose activity or activities are of interest. Taking cytotoxicity assays as an example, the reagent cocktail may be added to the cells under test before, after, or simultaneously with the potentially cytotoxic agent, depending on the kind of test being performed. If a quantitative determination of killing rate were desired, the cells could be mixed with the agent first and incubated for a fixed interval, after which the reagent cocktail would be added; this would provide an accurate picture of aggregate cell death over time. For maximum speed, reagent cocktail, cells, and the potentially cytotoxic agent could be mixed simultaneously; depending on the speed of killing, a signal could be obtained within minutes, or possibly even less than one minute. Finally, mixing the reagent cocktail with cells before addition of the potentially cytotoxic agent would allow comparison of the viability before and after treatment. These last two modes would also allow the user to follow the whole toxicity reaction in real time. A calibration standard of cells could be used to obtain absolute quantification. Note that the homogeneous nature of this aspect of the invention distinguishes it, in the case of cytotoxicity, from the GPL method, in which the assay reagents are not added in a single reagent mixture; instead the GPL method requires multiple transfers and incubations, first from the sample being tested to the xe2x80x9cGPxe2x80x9d cocktail; next, following an incubation, from the GP cocktail to the luciferase cocktail, which must also be aliquoted separately. Moreover, the GPL assay is not compatible with live cells, which must be separated by centrifugation, filtration, or another method before the first transfer. In the present invention, all constituents necessary for the assay are added in a single aliquot to the sample being tested, and there is no need to remove live cells from the supernatant.
In a related aspect, the present invention provides a set of methods for measuring cell proliferation. In this mode, the cells are killed by addition of a lytic agent before, after, or simultaneously with addition of the reagent cocktail. If the reagent cocktail is added before the lytic agent, a readout is obtained both of cells killed by processes under study (before addition of the lytic agent) and total cells present (after addition of the lytic agent). If the reagent cocktail is added after the lytic agent, a consistent increase or decrease in the luminance signal may be obtained, representing the total number of cells. If the lytic agent and reagent cocktail are added simultaneously, maximum throughput may be achieved, and the lytic process may be observed in real time (this is also true when the reagent cocktail is added first). Note that this feature also distinguishes the present invention from the GPL method. In the GPL assay, it is necessary to extract live cells from the sample being tested before addition of the GPL reagents, since in many cases these live cells could be killed by the GPL reagents. Thus failure to remove these live cells would lead to a mixed signal of actual cytotoxicity and a portion of the cells that were still alive prior to addition of the GPL reagents. The present invention does not suffer from this limitation, since the reagent mixture is compatible with all types of live cells that have been tested, including several mammalian cell lines, and Gram positive and Gram negative bacteria.
In a preferred embodiment, the present invention provides a set of methods for measuring cell proliferation and cytotoxicity in the same experiment, in a simple, two-step process which maintains the homogeneous nature of the assay. The reagent cocktail is added to cells before, during, or after initiation of the cytotoxic process. Following an incubation to obtain a luminance increase or decrease to obtain a cytotoxicity readout (typically 0.5 to 10 minutes), the lytic agent is added. The luminance increase or decrease following addition of the lytic agent represents the total biomass, alive and dead, at the time of the assay (proliferation readout). The live biomass (as of the time immediately before the lytic step) may generally be calculated by subtracting the toxicity readout from the total-biomass signal. The option of measuring both cytotoxicity and proliferation (or viability) in the same sample distinguishes the present invention from other available liquid-phase cytotoxicity and proliferation assays.
In another aspect, the present invention provides a set of methods and compositions for killing live cells of various types in a manner consistent with accurate reading of luminance due to enzyme release after the lytic step.
In another aspect, the present invention provides a set of methods and compositions for protecting an oxidation- and/or proteolysis-sensitive enzyme released by dying cells from oxidation and/or proteolysis during an initial incubation period, such that enzyme released by cells that die during the incubation period will be measurable at the end of that period.
In another aspect, the present invention provides a set of methods for detecting membrane damage, with or without associated cytotoxicity. Membrane damage associated with cell death is detected as cytotoxicity as discussed above. Membrane damage can also be detected separately from cell death (i.e., non-fatal damage) by performing assays by one of the specified methods for enzyme release, followed by an optional recovery phase and subsequently by a proliferation or viability assay, such as the CFU assay, a metabolism-based assay, or the proliferation mode of the present invention.
In another aspect, the present invention provides a set of methods of detecting enzymatic activity by coupling the enzymatic activity to production or consumption of a high-energy molecule that is a luciferase substrate.
In another aspect, the present invention provides a set of methods for optimizing a coupled luminescent assay for (1) time linearity, (2) linearity with enzyme or cell number to be measured, (3) compatibility with cells of various types, (4) homogeneous use, and (5) use in high-throughput screening (HTS).
In another aspect, the present invention provides a set of methods for optimizing the storage conditions of reaction cocktails and reaction cocktail ingredients with respect to physical form, storage of mixed or separate ingredients, temperature of storage, and time of storage.
In another aspect, the present invention provides a set of methods for automatic reduction of complex data by linear regression. These methods compute linear fits for all possible time ranges within a given data set and (1) report slopes and/or correlations for all ranges, and/or (2) select the time range or ranges with the highest correlation(s) and report these ranges, correlations, and slopes, and/or (3) select a time range or ranges with certain given fit characteristics from a given sample or set of samples (which could be calibration or other standards) and apply that range or those ranges to all or a subset of the remaining samples, and/or (4) detect and report exceptional aspects of data obtained from a given sample or samples, or from the entire run, such as poor signal strength, linearity, time correlation, or correlation with expected values, and/or (5) evaluate the characteristics of the run, such as linearity and/or signal strength, and either make an automated decision to stop or continue reading the samples or report the run characteristics to the user to allow the user to make that decision.
In another aspect, the present invention provides a set of methods of measuring cytotoxicity, membrane damage, cell proliferation, and enzymatic activity with the use of a stop reagent. The increase or decrease in the luminance signal is wholly or partially stopped by the reagent, allowing the user to treat the ending luminance value as the readout of the assay, such endpoint reading to take place at any time convenient to the user
In another aspect, the present invention provides a set of methods for HTS of compounds for any of a number of desirable or undesirable characteristics: (1) desirable cytotoxicity against an identified target, which may be a cancer cell type or an infectious microorganism; (2) undesirable cytotoxicity against normal cell types in a drug candidate; (3) growth-affecting characteristics; (4) membrane damage; or (5) inhibitory or rate-enhancing properties in a given enzymatic system. These methods involve preformulation of the reaction cocktail and preloading this cocktail into an injector of a luminometer, followed by homogeneous or non-homogeneous assay of the rate of increase or decrease in the luminance signal and either automated data reduction, non-automated data reduction, or the use of a stop reagent and a single readout. The HTS run may be (1) terminated after a single or fixed number of reads, (2) terminated automatically when certain criteria are achieved, or (3) terminated at the user""s discretion.
In another aspect, the present invention provides a set of methods for testing an individual patient""s cancer tumor cells or infecting microorganisms for sensitivity or resistance to a potential drug, drug mixture, or panel of drugs or drug mixtures.
In another aspect, the present invention provides a set of methods for detecting and quantifying apoptosis (programmed cell death). This may be accomplished as under the description of cytotoxicity measurement, above, or by coupled luminescent detection of the increase levels of nuclear G3PDH associated with apoptosis, or by a combination of these methods.
In another aspect, the present invention provides a set of methods for detecting the presence of live cells in environments that are intended to be sterile or have low bioburdens. This would be accomplished by taking a sample (either a liquid sample or a swabbed sample, which can then be transferred or washed into a liquid sample), using a lytic agent, and performing a coupled luminescent assay as described elsewhere under proliferation assays.
In another aspect, the present invention provides a set of methods for very sensitive detection of environmental toxins. This would be accomplished by mixing an environmental sample, such as an aliquot of seawater or residue from a wash of shellfish or other food samples, with a coupled luminescent reaction cocktail in the presence of a cell type known to be sensitive to the toxin in question, and measuring the resulting cytotoxicity.
In another aspect, the present invention provides a set of methods of detecting and/or quantifying free phosphate by coupling the presence of free phosphate to production of ATP via the activity of G3PDH and PGK, which are both supplied in the reagent mixture. Detection and/or quantification of free phosphate is of importance in biochemistry, enzymology, environmental science, and other areas.
In a second preferred embodiment, the present invention provides a set of methods for detecting the enzymatic activity of a phosphatase by quantifying the phosphate produced by the reaction of the phosphatase, which is accomplished by coupling the presence of free phosphate to production of ATP via the activity of G3PDH and PGK, which are both supplied in the reagent mixture. In this embodiment, the present invention enjoys a number of advantages over other phosphatase assays in current use, including great speed, extreme simplicity of operation, and the ability to use natural substrates, or, when they are unavailable, appropriately chosen phosphorylated peptide or protein substrates, or other phosphorylated molecules as similar as is practicable to the in vivo substrates.
In another aspect, the present invention provides a set of methods for detecting activity of intracellular phosphatases by optionally measuring phosphatase activity by the method described above before lysis, lysing the cells by one of the methods provided in the invention or by another method, and again measuring phosphatase activity. The principle may also be applied to measurement of phosphatase activity inside particular cellular organelles.
In another aspect, the present invention provides a set of methods for measuring activity of specific phosphatases, for which specific substrates are available, against a background of other phosphatases and/or free phosphate by measuring the quantity of phosphate present or the rate of phosphate production by the methods provided, adding the specific substrate or substrates, and again measuring the quantity of phosphate present (after a time interval of the user""s choice) or the rate of phosphate production.
These and other aspects of the present invention will become evident upon reference to the following detailed description and attached drawings.